Drugs can interfere with mitochondrial functions at various levels.1–10 For instance, they can inhibit mitochondrial respiration,2 DNA transcription, and glycolytic and fatty acid β-oxidation enzymatic activities.3 They may also induce the production of free radicals4 and a decrease in cellular endogenous antioxidants. Anesthetics, analgesics, and sedatives are among these drugs. In the mammalian heart, the inhalation anesthetic, isoflurane, inhibits complex I of the mitochondrial electron transport chain.11 In rat brain synaptosomes, propofol inhibits mitochondrial-coupled respiration and ATP production,12 and exposure of macrophages to propofol reduces Δψm and cellular ATP levels.13
Fentanyl is a widely used synthetic lipid-soluble short-acting narcotic analgesic.14–16 It is an opioid that interacts closely with μ-opioid receptors.17,18 It has been shown that fentanyl induces apoptosis of peripheral blood lymphocytes by disrupting Δψm and increasing production of reactive oxygen species.19 Vilela et al20 demonstrated that incubation of isolated brain mitochondria with fentanyl inhibits mitochondrial bioenergetics. Fentanyl is almost exclusively metabolized by the liver,18 and it reportedly impairs basal cellular oxygen consumption of isolated neonatal rat hepatocytes.21,22
Several studies have indicated that the opening of mitoKATP channels can modulate mitochondrial function.23–26 MitoKATP channels are located within the inner mitochondrial membrane and are involved in potassium influx into mitochondria. Under normal conditions, for proper mitochondrial function and cellular calcium homeostasis, the steady state of the mitochondrial matrix volume is maintained by a potassium efflux pathway (potassium-hydrogen antiporter).27 Potassium influx occurs by simple diffusion and mitoKATP channels.28–30 The opening of the mitoKATP channel will increase mitochondrial matrix volume.31,32 Transient-selective mitoKATP channel opening may provide functional protection against cellular injury and preserve cellular energy levels by regulating mitochondrial membrane swelling.29,33 However, continuous and prolonged mitoKATP channel opening induces cytochrome c release from mitochondrial intermembrane spaces, leading to apoptosis.29,33,34 Fentanyl enhances diazoxide-induced mitoKATP channel activity.35 In rats, fentanyl protects the heart against ischemic injury via opioid receptors and mitoKATP channel–linked mechanisms.36
Permeability glycoprotein (Pgp) is a transmembrane glycoprotein that functions as an ATP-dependent drug efflux pump and actively excretes a wide range of clinically important drugs and toxins out of the cell.37,38 Pgp can export structurally diverse hydrophobic compounds from the cell, and drugs that are transported by Pgp are identified as stimulators of its ATPase activity. In mice lacking Pgp, fentanyl-induced analgesia was increased and prolonged.39 Henthorn et al40 reported that fentanyl is a substrate of Pgp in cultured bovine brain microvascular endothelial cells. However, in a porcine kidney–derived cell line expressing human Pgp and transfected with the human multi-drug resistance 1 gene, fentanyl did not behave as a Pgp substrate.41 Thus, the effect of fentanyl on Pgp activity appears to be controversial in both in vitro and in vivo models.
The aim of the present study was to evaluate the effects of fentanyl on cellular and mitochondrial bioenergetics of cultured human hepatocytes and to investigate the potential contributing role of mitoKATP channels in this process. In addition, the in vitro effect of fentanyl on recombinant human Pgp ATPase activity in a cell membrane fraction was evaluated.
Chemicals and Reagents
Fentanyl citrate was obtained from Janssen-Cilag AG (Baar, Switzerland) and naloxone from OrPha Swiss GmbH (Küsnacht, Switzerland). All the reagents for cellular respiration, media for cell culture, and 5-hydroxydecanoate (5-HD) were obtained from Sigma-Aldrich (Buchs, Switzerland).
The human hepatoma cell line HepG2 (American Type Culture Collection) was cultured as described previously.42 All the experiments were performed with cells from the same batch at different passage numbers.
Incubation With Drugs
Quiescent cells were exposed at clinical and therapeutic blood concentrations (0.5 and 2 ng/mL)43–46 or higher (10 and 50 ng/mL) for 1 hour or pretreated with naloxone at 200 and 1000 ng/mL or 5-HD at 50 μM for 30 minutes, followed by incubation with fentanyl at 2 ng/mL for an additional hour at 37°C in a humid atmosphere (5% CO2, 95% air).
Cellular Respiration (High-Resolution Respirometry): Permeabilized and Intact Cells
After incubation, the cells were permeabilized to allow the entry of exogenous adenosine diphosphate (ADP) and oxidizable mitochondrial substrates to feed electrons into complexes of the respiratory system (complex I–dependent, complex II–dependent [after rotenone addition], and complex IV–dependent [after antimycin A addition] respiration). In addition, basal, coupled, and uncoupled respiration of intact cells was measured. This method evaluates oxygen consumption of intact cells without the addition of exogenous substrates and ADP, reproducing the respiratory function in the integrated cell. These methods are described in detail in the Supplemental Digital Content 1.
Measurement of Mitochondrial Membrane Potential (Δψm)
Measurement of Δψm in intact cells was performed using the dyes 5,5′,6′,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolocarbocyanine iodide (JC-1) and tetramethylrhodamine methyl ester (TMRM). These methods are described in detail in the Supplemental Digital Content 2 (http://links.lww.com/AA/B401).
Measurement of the HepG2 Cells’ ATP Content, ADP/ATP Ratio, Flavin-Adenine Dinucleotide Levels and ATP Synthase Enzymatic Activity, Reactive Oxygen Species/Reactive Nitrogen Species (ROS/RNS), and Mitochondrial Calcium Levels in HepG2 Cells
For these assessments, commercially available kits were used. These methods are described in detail in the Supplemental Digital Content 3 (http://links.lww.com/AA/B402).
Measurements of Extracellular L-Lactate Levels (Lactate Released to the Culture Medium)
Lactate concentrations in the cell culture supernatants were detected using a 96-well fluorescence-based assay kit. The method is described in the Supplemental Digital Content 4 (http://links.lww.com/AA/B403).
Transmission Electron Microscopy
Transmission electron microscopy was performed as described previously.42
Measurement of HepG2 Cells’ ATPase Activity and Cell-Free In Vitro Fentanyl-Stimulated Pgp ATPase Activity
The ATPase and Pgp ATPase activities of HepG2 cells were measured using commercially available kits (Supplemental Digital Content 5, http://links.lww.com/AA/B404).
The SPSS 21.0 software package (SPSS Inc, Chicago, IL) was used for statistical analysis. Based on our previous findings,47 we set n = 20 per experiment for the initially planned investigations (mitochondrial respiration). To explore potential mechanisms for and consequences of the obtained results, we used sample sizes between 8 and 32, unless performed on 96-well plates, where 24 to 76 experiments were conducted, based on the available space. For the Pgp ATPase activity, 4 experiments per group were performed as recommended by the manufacturer. Only in one investigation were further experiments added after a first statistical analysis (mitochondrial respiration at a fentanyl concentration of 50 ng/mL). In all other experiments, the sample number was fixed before statistical analysis. Because of the relatively low sample sizes in many of the experiments, and after statistical consultation, nonparametric tests were used for all analyses. Comparisons of the slopes of oxygen concentrations (cellular respiration) of each individual experiment (active compound and control) were made using the Wilcoxon matched-pairs signed rank test. For unpaired data (mitochondrial calcium content, ROS/RNS, flavin-adenine dinucleotide [FAD] levels, ADP/ATP ratio, cellular ATPase activity, and lactate levels) the Mann-Whitney test was used. Statistical analysis of mitochondrial ATP synthase enzymatic activity, membrane potential, total cellular ATP content, and Pgp ATPase activities were performed using a Kruskal-Wallis rank sum test, followed by Dunn multiple comparisons. Data are shown as median and interquartile range (IQR). Because of the multiple experiments performed, the addition of experiments after a first statistical analysis for mitochondrial respiration at a fentanyl concentration of 50 ng/mL, and after statistical advice, a conservative approach was taken: results with a P value <0.01 were considered significant and results with a P value >0.15 not significant. Data are indicated as median and IQR in text and figures and displayed as box plots in figures, where boxes indicate median and IQR and whiskers represent minimum and maximum.
Cell and Mitochondrial Morphology Is Not Affected by Fentanyl
To evaluate whether fentanyl induces alterations in cellular and mitochondrial morphology, electron microscopy was performed. Transmission electron microscopic examination revealed no major alterations in cellular or mitochondrial ultrastructure under fentanyl treatment (n = 3) (Figure 1).
Respiratory Function in the Integrated Cell Using Endogenous Substrates Is Not Affected by Fentanyl
To evaluate whether fentanyl stimulation is implicated in hepatic mitochondrial dysfunction, we first investigated the effects of fentanyl exposure on respiratory function in the integrated cell using endogenous substrates and measured maximal capacity of the electron transport chain using carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (FCCP). Compared with our previous investigations of effects of remifentanyl on hepatocytes’ mitochondrial respiration,47 we used a slightly higher number of experiments (20 per condition). A representative diagram of measurement of respiration rates in intact HepG2 using high-resolution respirometry is shown in Figure 2A. The uncoupler FCCP was titrated to an optimum concentration for maximum stimulation of flux. In the absence of exogenous substrates and ADP, fentanyl (2 ng/mL, 1-hour incubation) did not affect basal cellular respiration (Z: 0.000, P = 1.000), oligomycin-insensitive (nonphosphorylating respiration; Z: −0.751, P = 0.452), oligomycin-sensitive (ATP turnover; Z: −0.228, P = 0.819), or FCCP-uncoupled maximal respiration rates (Z: −0.280, P = 0.779; Figure 2B; n = 20 each). Uncoupled respiratory control ratios (the ratio between FCCP and oligomycin-insensitive respiration rates; median [IQR]; control: 8.1 [7.3–8.9]; fentanyl: 9.0 [7.6–9.9]; Z: −1.868, P = 0.062) may have been affected, but coupling efficiency (the ratio between oligomycin-sensitive and basal respiration rates; Z: −0.787, P = 0.431) was not (Figure 2C; n = 20 each).
Cellular Oxygen Consumption Is Impaired After Incubation With Fentanyl Using Exogenous Substrates Specific for Mitochondrial Complexes I, II, and IV
We further investigated whether fentanyl exposure induces alteration in mitochondrial respiration using exogenous substrates (maximal stimulated respiration; n = 20 per condition). A representative diagram of measurement of respiration rates in digitonin-permeabilized HepG2 cells using high-resolution respirometry is shown in Figure 3A. Fentanyl at 0.5 ng/mL did not reduce mitochondrial (complex I–dependent [Z: −0.655, P = 0.513] and complex II–dependent respiration [Z: −0.579, P = 0.562]). The effect on complex IV–dependent respiration was not large enough to be significant (control: 78.0 [62.5–89.8]; fentanyl: 70.0 [64.0–76.0] pmol/[seconds × million cells], Z: −1.946, P = 0.052; n = 20 each; Figure 3B). Fentanyl at 2 ng/mL induced a significant reduction in complex IV–dependent respiration (control: 90.0 [78.3–106.0]; fentanyl: 79.0 [68.3–89.5] pmol/[seconds × million cells]; Z: −3.305, P = 0.001), but probably not in complex I–dependent respiration (control: 51.0 [47.0–64.0]; fentanyl: 48.0 [44.3–52.0] pmol/[seconds × million cells]; Z: −1.552, P = 0.121). The effect on complex II–dependent respiration was not large enough to be significant (control: 103.5 [81.3–118.0]; fentanyl: 93.5 [73.5–99.0] pmol/[seconds × million cells]; Z: −2.316, P = 0.021; Figure 3C; n = 20 each). Fentanyl at 10 ng/mL did not significantly impair respiration of complex I–dependent (Z: −0.935, P = 0.350) and complex II–dependent respiration (Z: −0.411, P = 0.681), and the effect on complex IV–dependent respiration was not large enough to be significant (control: 83.0 [71.5–104.5]; fentanyl: 75.5 [65.5–89.5] pmol/[seconds × million cells]; Z: −2.223, P = 0.026; Figure 3D; n = 20 each). The effects of fentanyl at 50 ng/mL were at the limit of statistical significance after 20 experiments. Therefore, 10 experiments per condition were added. This resulted in a reduction in complex I–dependent (control: 35.9 [29.0–43.0]; fentanyl: 32.1 [25.0–39.2] pmol/[seconds × million cells]; Z: −2.985, P = 0.003), complex II–dependent (control: 67.0 [54.4–75.1]; fentanyl: 54.5 [45.8–67.3] pmol/[seconds × million cells]; Z: −3.264, P = 0.001), and complex IV–dependent respiration (control: 68.0 [62.4–77.5]; fentanyl: 60.0 [52.8–70.2] pmol/[seconds × million cells]; Z: −3.094, P = 0.002; Figure 3E, n = 30 each). Citric acid, the inactive ingredient of fentanyl, did not affect cellular respiration (data not shown).
Fentanyl Reduces Respiration Efficiency When Exogenous Fatty Acid Palmitate Is Used as a Substrate
In our experiments, the effect of fentanyl on stimulated mitochondrial respiration was in the magnitude of 10% to 20%. In studies in hepatocytes from suckling rats, morphine inhibited oxygen consumption by up to 25% and fentanyl by up to 40%21 when palmitate was used as a substrate for mitochondrial respiration. To evaluate whether fentanyl affects mitochondrial respiration to a greater extent when exogenous fatty acids are used as a substrate, we evaluated the effect of fentanyl on palmitate-dependent respiration. Fentanyl at 50 ng/mL reduced the efficiency of mitochondrial respiration (control: 5.8 [4.8–7.1]; fentanyl: 4.7 [4.4–5.2]; Z: −2.654, P = 0.008; n = 10; Figure 4C). The reduction at the other concentrations (fentanyl 2 ng/mL: 5.4 [5.05–5.9]; fentanyl: 4.5 [4.1–5.0]; Z: −2.449, P = 0.012; fentanyl 10 ng/mL: control: 5.7 [5.1–6.1]; fentanyl: 4.6 [4.0–5.1]; Z: −1.990, P = 0.047; n = 10 each; Figure 4, A and B) was not large enough to be significant. Neither dose affected state 3 respiration significantly (fentanyl 2 ng/mL: Z: −1.020, P = 0.308; fentanyl 10 ng/mL: Z: −0.255, P = 0.799; fentanyl 50 ng/mL: Z: −0.764, P = 0.445; n = 10 each; Figure 4, A–C). The effect on state 4o was not large enough to be significant (fentanyl 50 ng/mL: control: 5.1 [4.0–5.7]; fentanyl: 6.5 [4.9–7.6] pmol/[seconds × million cells]; Z: −2.142, P = 0.032; Figure 4, A–C).
All effects of palmitate disappeared if cells were pretreated with 5-HD (50 μM, 30 minutes) before the addition of fentanyl (2 ng/mL, 1 hour; state 3: Z: −0.534, P = 0.594; state 4o: Z: −0.474, P = 0.635; respiratory control ratio calculated as state 3/state 4o.: Z: −1.192, P = 0.233; n = 9 each; Figure 4D).
Fentanyl-Induced Reduction in Cellular Oxygen Consumption Is Prevented by Preincubation With Naloxone
We further evaluated whether antagonism with naloxone (an opioid receptor antagonist) can prevent fentanyl-induced reduction in cellular oxygen consumption. Cells were preincubated with naloxone for 1.5 hours either alone (Figure 5, B and D) or preincubated with naloxone (200 and 1000 ng/mL) for 30 minutes before the addition of fentanyl (2 ng/mL, 1-hour incubation; n = 20 each) (Figure 5, A and C). Controls for each series were incubated with medium alone. Antagonism with naloxone at 1000 ng/mL abolished the effect of fentanyl: naloxone 1000 ng/mL, complex I, Z: −0.981, P = 0.326; complex II, Z: −0.282, P = 0.778; complex IV, Z: −0.907, P = 0.364; n = 20 each; Figure 5, A and C). Naloxone at 200 ng/mL abolished the effect of fentanyl on complex I, Z: −0.841, P = 0.400, and was likely to abolish the effects on complex II (control: 76.5 [59.6–104.5]; fentanyl + naloxone: 71.5 [60.3–87.2] pmol/[seconds × million cells]; Z: −1.609, P = 0.108) and complex IV (control: 77.5 [68.0–102.0]; fentanyl + naloxone: 77.5 [65.0–85.8] pmol/[seconds × million cells]; Z: −1.681, P = 0.093).
Incubation of the cells with naloxone alone at nanogram per milliliter had no significant effect (naloxone 1000 g/mL, complex 1, Z: −1.252, P = 0.211; complex II, Z: −0.131, P = 0.896; complex IV, Z: −0.168, P = 0.867; n = 20 each; Figure 5, B and D). Incubation of the cells with naloxone at 200 ng/mL had no significant effect either on complex II (Z: −1.410, P = 0.159) or on complex IV (Z: −0.897, P = 0.370). The effect on complex I was not large enough to be significant (naloxone 200 ng/mL, control: 41.0 [38.0–45.0]; naloxone: 47.5 [42.3–53.0] pmol/[seconds × million cells]; Z: −2.207, P = 0.027).
Fentanyl-Induced Reduction in Cellular Respiration Is Abolished by 5-HD (an Inhibitor of Mitochondrial ATP-Sensitive Potassium [mitoKATP] Channels)
Because fentanyl enhances the effects of mitoKATP channel–opening drugs,35 we further evaluated whether 5-HD abolishes fentanyl-induced reduction in cellular respiration. Cells pretreated with the mitoKATP channel inhibitor 5-HD (50 μM, 30 minutes) before the addition of fentanyl (2 ng/mL, 1 hour) exhibited no significant changes in complex activities in comparison with controls (complex I, Z: −1.008, P = 0.313; complex II, Z: −1.269, P = 0.204; complex IV, Z: −1.344, P = 0.179; n = 20 each; Figure 6A). Incubation of the cells with 5-HD alone did not affect cellular respiration (complex I, Z: −0.859, P = 0.391; complex II, Z: −0.448, P = 0.654; complex IV, Z: −0.859, P = 0.391; n = 20 each; Figure 6B).
Mitochondrial Membrane Potential (Δψm) Is Not Affected by Fentanyl Using the Dyes TMRM and JC-1
Because reduced and inefficient mitochondrial respiration can result from lowered Δψm, we next investigated whether fentanyl induces alterations in mitochondrial membrane potential. For these experiments, we used 2 different dyes, JC-1 and TMRM (Figure 7 and 8). Fentanyl at 2 ng/mL for 1 hour did not induce a significant reduction in mitochondrial membrane potential measured with TMRM (control: 43.2 [34.9–56.3]; fentanyl: 44.2 [36.4–52.8]; 5-HD: 43.9 [33.5–55.7]; fentanyl + 5-HD: 46.8 [36.7–57.3] relative fluorescence units per nanogram cellular protein; Kruskal-Wallis rank sum test:
, P = 0.868; n = 45 each; Figure 7) and also did not appear to induce a significant reduction if measured with JC-1 (control: 0.43 [0.40–0.54] [n = 73]; fentanyl: 0.43 [0.38–0.47] [n = 74]; fentanyl + 5-HD: 0.44 [0.39–0.49] 590/530 fluorescence ratio [n = 76]; Kruskal-Wallis rank sum test:
, P = 0.094; Figure 8).
Cellular ATP Content but Not ATP Synthase Activity (Complex V or F1F0 ATPase) Is Reduced After Incubation With Fentanyl
Because [mitoKATP] channel stimulation and reduced mitochondrial respiration can decrease ATP content, this was evaluated in the next experiment. Although fentanyl at 2 ng/mL for 1 hour reduced cellular ATP content, this effect was not significantly attenuated by preincubation with 5-HD (50 μM) (control: 6.1 [5.3–7.4]; fentanyl: 3.6 [2.7–4.2]; fentanyl + 5-HD: 4.0 [3.4–4.8]; Kruskal-Wallis rank sum test:
, P < 0.001, followed by Dunn multiple comparisons test: control versus fentanyl: P < 0.001, fentanyl versus 5-HD + fentanyl: P = 0.126; n = 54 each; Figure 9A). The effect of fentanyl at 2 ng/mL for 1 hour was not large enough to alter cellular ATP synthase activity (control: 4.3 [3.9–4.7] mM/min/mg cellular protein; fentanyl: 4.1 [3.9–4.2] mM/min/mg cellular protein; fentanyl + 5-HD: 4.5 [4.0–4.9] mM/min/mg cellular protein; Kruskal-Wallis rank sum test:
, P =0.081; Figure 9B).
The ATP synthase inhibitor, oligomycin, at 500 nM inhibited the activity of the F1FO ATPase by >90% (data not shown), indicating the specificity of the reaction to the F1FO ATPase complex.
Lactate Released to the Culture Medium Is Not Affected by Fentanyl
To evaluate the significance of the decreased ATP content, we evaluated whether it was compensated for by increased glycolysis, which should increase lactate concentration in the cell culture medium. Incubation of the cells with fentanyl for 1 hour at 2 ng/mL did not significantly affect extracellular lactate levels (control: 1.8 [1.6–2.3] μM; fentanyl: 2.9 [2.2–3.8] μM; 5-HD: 3.3 [2.7–3.4] μM; fentanyl + 5-HD: 2.5 [2.4–3.8] μM; Kruskal-Wallis rank sum test:
; P = 0.032; n = 6 each; Figure 10).
ATPase Activity and ADP/ATP Ratio Are Not Affected by Exposure to Fentanyl
To characterize alterations in ATP metabolism suggested by the reduced ATP content, the activity of mitochondrial ATP synthase and the ADP/ATP ratio were studied. Treatment of the cells with fentanyl at 2 ng/mL for 1 hour neither induced an increase in total cellular ATPase activity (Mann-Whitney U test: Z: −1.172, P = 0.247; n = 10 each; Figure 11A) nor changed the ADP/ATP ratio (Mann-Whitney U test: Z: −0.988, P = 0.327; n = 20 each; Figure 11B).
Fentanyl Stimulates Cell-Free In Vitro Pgp ATPase Activity
5-HD did not seem to abolish fentanyl-induced reduction in cellular ATP content, and total ATPase activity of cells treated with fentanyl was unchanged. Measurements of total cellular ATPase activity can be challenging because each specific cellular ATPase may need specific conditions for optimal ATPase activation.48,49 Thus, we cannot exclude the possibility that there were masked changes in specific cellular ATPase activities under our assay conditions. Because it has been suggested that drugs such as morphine,50 fentanyl,40 and verapamil51 are substrates of Pgp, which is an ATP-dependent cellular transport or efflux ATPase, we next investigated whether fentanyl affects Pgp ATPase activity.37,38 For these experiments, recombinant human Pgp membrane fractions were treated with fentanyl at 2, 10, and 40 ng/mL and excess ATP for 30 minutes, and ATP consumption rates were measured. Fentanyl stimulated the rate of ATP consumption similarly to the positive control verapamil (Kruskal-Wallis rank sum test:
, P = 0.004; n = 4 each, according to the Pgp-Glo™ Assay System manufacturer’s instructions [Promega, Madison, WI]; Figure 12) and as found by others.40
Fentanyl Reduces Cellular ATP Content in a Dose-Dependent Manner
Because Pgp ATPase activity demonstrated dose responsiveness to fentanyl, additional experiments were performed to determine the cellular ATP content with escalating doses of fentanyl. For these experiments, cells were treated with fentanyl at 0.5, 2, 10, and 50 ng/mL for 1 hour, and cellular ATP content was measured. Fentanyl reduced cellular ATP content in a dose-dependent manner (Kruskal-Wallis rank sum test:
, P < 0.001; n = 24 each; Figure 13).
Mitochondrial Calcium Content, ROS/RNS, and FAD Levels
Finally, to elucidate further mechanisms and consequences of fentanyl-induced impairment of mitochondrial respiration and decreased ATP content—potentially, but not necessarily, related to mitoKATP channel stimulation—we measured intramitochondrial calcium concentration and ROS/RNS and FAD levels.
Because an excessive mitochondrial calcium level can impair respiratory capacity,52 we first investigated mitochondrial calcium content. Fentanyl had no effect on intramitochondrial calcium concentration (Mann-Whitney U test: Z: −1.116, P = 0.265; n = 32/16 [controls/fentanyl]; Figure 14A). Inefficiency in the mitochondrial respiratory capacity and ATP production may induce alterations in mitochondrial ROS generation. Depending on the condition, mitochondrial ROS generation can increase or decrease.53–55 For example, increased mitochondrial ROS generation has been shown when mitochondrial proton motive force increases,54 whereas, under low oxygen levels, impaired ROS generation has been reported.55 Therefore, we further evaluated cellular ROS/RNS levels. Fentanyl did not alter ROS/RNS levels significantly (control: 4.6 [4.3–4.8] vs fentanyl: 4.3 [4.1–4.6] nM/μg; Mann-Whitney U test: Z: −2.083, P = 0.037; n = 24 each; Figure 14B). To further evaluate and correlate mitochondrial electron transport chain substrate availability to respiration, we measured mitochondrial FAD levels. The effect of fentanyl cellular FAD levels was not large enough to be significant (control: 0.11 [0.10–0.11] vs fentanyl: 0.10 [0.10–0.11] nM/μg; Mann-Whitney U test: Z: −2.205, P = 0.028, n = 8 each; Figure 14C).
In the present study, we demonstrate that fentanyl slightly reduces cultured human hepatocytes’ mitochondrial respiration by a mechanism that is blocked by a mitoKATP channel antagonist. 5-HD seems to abolish a fentanyl-induced reduction in cellular ATP content only marginally if at all. No major visible ultrastructural alterations were found after incubation with fentanyl.
We measured mitochondrial respiration using specific exogenous substrates for mitochondrial complexes I, II, and IV, and, in addition, maximal respiration due to oxidation of the exogenous fatty acid palmitate. Therapeutic blood levels of fentanyl43–46 decreased complex II–dependent and complex IV–dependent mitochondrial and palmitate-dependent respiration although the effect was small.
In 2 studies, in neonatal rat hepatocytes, it has already been shown that analgesic doses of fentanyl reduced palmitate-dependent cellular oxygen consumption.21,22
In our study, naloxone, which is an opioid inverse agonist,17,18 attenuated fentanyl-induced reduction in mitochondrial respiration, indicating that the fentanyl effect was mediated by an opioid receptor mechanism. We incubated the cells with fentanyl for only 1 hour because fentanyl has a rapid onset and short duration of action, with a plasma half-life of 90 minutes and an elimination half-time of a few hours.56
The mitochondrial respiration impairment was accompanied by reductions in cellular ATP content. The reduction in ATP levels in the presence of fentanyl may be due to an effect of the drug on oxygen consumption or ATP synthase activity or on other components of mitochondrial/cellular function. In isolated rat brain mitochondria, high concentrations of fentanyl interfered with the mitochondrial electron transport chain (complexes II and IV).20 Reduced mitochondrial respiration and ATP production may induce the acceleration of glycolysis and lactate production to provide additional ATP. However, we found no increase in extracellular lactate levels with fentanyl treatment. Surprisingly, despite a reduction in mitochondrial ATP content, the ADP/ATP ratio remained unchanged after fentanyl exposure. Although the kit we used for total ATP concentration is more sensitive and inactivates ATPases in contrast to the kit for the assessment of the ADP/ATP ratio, we suggest that decreased ATP availability due to fentanyl-induced impairment of mitochondrial respiration, combined with increased ATP use for the Pgp efflux pump, can result in a decrease in total adenosine store and thereby cause a decrease in both ATP and ADP.
Fentanyl did not induce a reduction in mitochondrial membrane potential, as measured with TMRM, and also did not appear to induce a significant reduction if measured with JC-1. Changes in the mitochondrial respiratory capacity and ATP production may alter (increase or decrease) mitochondrial ROS generation.53,54 Increased mitochondrial ROS generation has been shown when mitochondrial proton motive force increases,54 and under hypoxic conditions, ROS generation has been reported to be decreased.55 In 1 study, incubation of peripheral blood lymphocytes with fentanyl induced apoptosis with disrupted Δψm and increased production of ROS.19 In our study in HepG2 cells, extracellular ROS concentrations were not altered significantly, and we did not find evidence that fentanyl uncoupled mitochondrial respiration.
MitoKATP channels may contribute to the regulation of mitochondrial function,23–26,57 and the presence of mitoKATP channels in hepatocytes has been shown.58 In rabbit ventricular myocytes, diazoxide, a mitoKATP channel opener, induced reversible oxidation of flavoproteins (an index of mitochondrial redox state).59 In isolated cardiac mitochondria, mitoKATP channel openers decreased mitochondrial membrane potential and ATP production and produced swelling.60 It has been suggested that potassium influx into the mitochondrial matrix increases matrix volume59 and reduces the potential across the inner mitochondrial membrane.30
In our present study, 5-HD (a mitoKATP channel antagonist) did not seem to abolish fentanyl-induced reduction in cellular ATP content. An alternative explanation for the reduction of cellular ATP content is the activation of cellular ATPase activity. Therefore, we first investigated whether fentanyl increases total cellular ATPase activity. The effect of fentanyl was not large enough to significantly alter total cellular ATPase activity. However, measurement of total cellular ATPase activity is challenging because each specific cellular ATPase may have different critical conditions for optimal ATPase activation.48,49 Thus, it is possible that there are masked specific cellular ATPase activities under our assay conditions. We then investigated whether fentanyl increases a specific cellular ATPase activity. It has been reported that opioids are Pgp substrates.61–63 Drugs, such as morphine, which stimulate Pgp ATPase activity are usually compounds for cellular transport or efflux.50 However, the effect of fentanyl on Pgp activity is somewhat controversial.39–41 In an in vitro study assessing transcellular movement of fentanyl in porcine kidney epithelial cells expressing human Pgp,41 fentanyl did not behave as a Pgp substrate. In contrast, Henthorn et al40 reported that, in cultured bovine brain microvascular endothelial cells, fentanyl is a Pgp substrate. The 2 different cell types used in these studies might have differed in terms of Pgp expression and activity, as well as in cell membrane composition and characteristics.40,41 In addition, different methods have been used to evaluate fentanyl as a substrate for Pgp. In an in vivo study in mice lacking Pgp, fentanyl-induced analgesia was increased and prolonged, suggesting that Pgp alters the efficiency of fentanyl.39 However, the authors did not perform detailed pharmacokinetic studies for fentanyl in plasma, and the effect of Pgp on fentanyl concentration in the brain was not evaluated.
In our present study using an in vitro cell-free system assay, we were able to demonstrate that fentanyl increases Pgp ATPase activity, suggesting that it is indeed a Pgp substrate. Protein expression of Pgp in HepG2 cells has been reported,64 and increased Pgp ATPase activity upon fentanyl treatment might account for the observed reduced ATP levels in our experiments. Because the Pgp ATPase activity demonstrated a dose responsiveness to fentanyl, we evaluated the effect of escalating fentanyl doses on cellular ATP content. We were able to demonstrate a dose-dependent reduction in ATP content. We suggest that this represents an increased need for ATP to power the Pgp efflux pump as the fentanyl concentration is increased. Our combined data indicate that ATP content is affected at fentanyl concentrations between 2 and 10 ng/mL, probably also dependent on methodological factors such as type of plates used to culture cells and the cell passage number.
Schematic representation of the proposed effect of fentanyl on mitochondrial function and Pgp activity is shown in Figure 15. μ-Opioid receptors are transmembrane proteins that couple to inhibitory G-proteins.65 After activation of μ-opioid receptors by fentanyl, the Gα and Gβγ subunits dissociate from one another and subsequently act on intracellular effector pathways such as tyrosine kinase and protein kinase C (PKC) pathways.65–67 It has been suggested that PKC activators, such as phorbol esters, enhance the opening of mitoKATP channels,68,69 and the opening of the mitoKATP channel by diazoxide (a mitoKATP channel opener) is mediated by the PKC signaling pathway.70
In our study, fentanyl reduced cultured human hepatocytes’ mitochondrial respiration by a mechanism that was blocked by a mitoKATP channel antagonist (5-HD). Opening of the mitoKATP channel was probably mediated by the activation of the PKC pathway. Antagonism with naloxone abolished a fentanyl-induced decrease in cellular respiration, indicating that fentanyl exerts these pharmacologic actions via receptor-mediated mechanisms. Fentanyl is highly lipid soluble and can pass across the cell membranes. 5-HD could not prevent fentanyl-induced reduction in cellular ATP content, indicating activation of cellular ATPases. Using an in vitro assay, we show that fentanyl is a substrate for efflux pump Pgp and directly stimulates its ATPase activity. In intact cells, PKC activators have been shown to increase phosphorylation and activity of the Pgp71–74 and decrease cellular drug accumulation.73,75,76 Therefore, fentanyl might exert its effect on the Pgp efflux pump either by direct interaction with it, by μ-opioid receptor–mediated activation of the PKC pathway or by a combination of both.
A limitation of the present study is the use of human hepatoma cell line HepG2. Because HepG2 cells contain high levels of mitochondria and mitochondrial DNA content, they are an excellent choice to study mitochondrial toxicity.77 However, HepG2 shows a different phenotype and certain dissimilarities when compared with primary human hepatocytes.78,79 For example, HepG2 cells secrete lower amounts of triglycerides78 and contain low levels of cytochrome P450 enzymes, which are involved in drug metabolism.80 Therefore, further experiments are warranted with primary hepatocytes or whole organs, before the results of our study can be translated to the human condition.
Although the effects of fentanyl on mitochondrial function that we documented are rather small, opioids—especially at high doses—can cause liver failure. As an example, the agonist/antagonist opioid buprenorphine, if injected IV, results in 80 times higher peak concentrations than after sublingual administration.81 This has been associated with cytolytic hepatitis.82 The mechanism related to this was evaluated in rat hepatocytes and in living mice, where high doses of buprenorphin were associated with impaired mitochondrial respiration and ATP formation.82 Similarly, street heroin, composed of heroin, 6-monoacetylmorphine, and morphine, induced mitochondrial dysfunction and apoptosis in cortical neuron cells.83 In rat liver mitochondria, morphine decreased mitochondrial ATPase, activity,84 and in SH-SY5Y cells, methadone caused concentration-dependent depletion of ATP levels.85
In patients receiving fentanyl for pain control, plasma concentrations are certainly lower than in drugs addicts. Nevertheless, in patients treated with fentanyl after cardiac arrest, fentanyl blood concentrations of up to 18 ng/mL were recorded,44 and in patients with cancer exposed to transdermal fentanyl, plasma concentrations of up to 14.7 ng/mL have been documented.45,46 Fentanyl plasma concentrations were highest in cachectic patients and in those with reduced renal function, signs of liver injury and/or inflammation.46 Therefore, our results suggest that mitochondrial dysfunction may play a significant role in patients exposed to very high fentanyl doses, such as in drug addicts and cachectic patients with liver and/or renal failure and infection. It has been proposed that, in the latter group of patients, plasma fentanyl concentrations could be measured and fentanyl administration reduced if the concentration is high.46 It is not known whether patients with intensive care unit-acquired weakness, significant muscle loss, organ failure, and inflammation achieve similarly high or possibly higher fentanyl concentrations. If these patients develop energetic failure, drug-induced mitochondrial dysfunction may be considered.
In summary, our data suggest that fentanyl reduces stimulated cultured human hepatocytes’ mitochondrial respiration by a mechanism that is blocked by a mitoKATP channel antagonist. 5-HD does not seem to completely abolish a fentanyl-induced reduction in cellular ATP content. This could not be explained by alterations in total cellular ATPase activity; however, fentanyl markedly stimulated the in vitro ATPase activity of recombinant human Pgp.
Name: Siamak Djafarzadeh, PhD.
Contribution: This author helped design the study, conduct the study, collect the data, analyze the data, and prepare the manuscript.
Name: Madhusudanarao Vuda, PhD.
Contribution: This author helped conduct the study, collect the data, and analyze the data.
Name: Victor Jeger, MD, PhD.
Contribution: This author helped conduct the study, collect the data, and analyze the data.
Name: Jukka Takala, MD, PhD.
Contribution: This author helped design the study, analyze the data, and prepare the manuscript.
Name: Stephan M. Jakob, MD, PhD.
Contribution: This author helped design the study, conduct the study, collect the data, analyze the data, and prepare the manuscript.
This manuscript was handled by: Markus W. Hollmann, MD, PhD, DEAA.
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